Abstract

The regulation of higher-order chromatin structure is complex and dynamic, and a full understanding of the suite of mechanisms governing this architecture is lacking. Here, we reveal the noncanonical SMC protein Smchd1 to be a novel regulator of long-range chromatin interactions in mice, and we add Smchd1 to the canon of epigenetic proteins required for Hox-gene regulation. The effect of losing Smchd1-dependent chromatin interactions has varying outcomes that depend on chromatin context. At autosomal targets transcriptionally sensitive to Smchd1 deletion, we found increased short-range interactions and ectopic enhancer activation. In contrast, the inactive X chromosome was transcriptionally refractive to Smchd1 ablation, despite chromosome-wide increases in short-range interactions. In the inactive X, we observed spreading of trimethylated histone H3 K27 (H3K27me3) domains into regions not normally decorated by this mark. Together, these data suggest that Smchd1 is able to insulate chromatin, thereby limiting access to other chromatin-modifying proteins.

Access optionsAccess options

Rent or Buy article

Get time limited or full article access on ReadCube.

from$8.99

All prices are NET prices.

Additional information

Publisher’s note: Springer Nature remains neutral with regard to jurisdictional claims in published maps and institutional affiliations.

References

  1. 1.

    Bouwman, B. A. & de Laat, W. Getting the genome in shape: the formation of loops, domains and compartments. Genome Biol. 16, 154 (2015).

  2. 2.

    Dekker, J. & Mirny, L. The 3D genome as moderator of chromosomal communication. Cell 164, 1110–1121 (2016).

  3. 3.

    Kakui, Y. & Uhlmann, F. SMC complexes orchestrate the mitotic chromatin interaction landscape. Curr. Genet. 64, 335–339 (2018).

  4. 4.

    Lieberman-Aiden, E. et al. Comprehensive mapping of long-range interactions reveals folding principles of the human genome. Science 326, 289–293 (2009).

  5. 5.

    Rao, S. S. P. et al. A 3D map of the human genome at kilobase resolution reveals principles of chromatin looping. Cell 159, 1665–1680 (2014).

  6. 6.

    Schmitt, A. D., Hu, M. & Ren, B. Genome-wide mapping and analysis of chromosome architecture. Nat. Rev. Mol. Cell Biol. 17, 743–755 (2016).

  7. 7.

    Dixon, J. R. et al. Topological domains in mammalian genomes identified by analysis of chromatin interactions. Nature 485, 376–380 (2012).

  8. 8.

    Nora, E. P. et al. Targeted degradation of CTCF decouples local insulation of chromosome domains from genomic compartmentalization. Cell 169, 930–944.e22 (2017).

  9. 9.

    Schwarzer, W. et al. Two independent modes of chromatin organization revealed by cohesin removal. Nature 551, 51–56 (2017).

  10. 10.

    Eskeland, R. et al. Ring1B compacts chromatin structure and represses gene expression independent of histone ubiquitination. Mol. Cell 38, 452–464 (2010).

  11. 11.

    Kundu, S. et al. Polycomb repressive complex 1 generates discrete compacted domains that change during differentiation. Mol. Cell 65, 432–446.e5 (2017).

  12. 12.

    Blewitt, M. E. et al. SmcHD1, containing a structural-maintenance-of-chromosomes hinge domain, has a critical role in X inactivation. Nat. Genet. 40, 663–669 (2008).

  13. 13.

    Chiu, A., Revenkova, E. & Jessberger, R. DNA interaction and dimerization of eukaryotic SMC hinge domains. J. Biol. Chem. 279, 26233–26242 (2004).

  14. 14.

    Haering, C. H., Löwe, J., Hochwagen, A. & Nasmyth, K. Molecular architecture of SMC proteins and the yeast cohesin complex. Mol. Cell 9, 773–788 (2002).

  15. 15.

    Hirano, M. & Hirano, T. Hinge-mediated dimerization of SMC protein is essential for its dynamic interaction with DNA. EMBO J. 21, 5733–5744 (2002).

  16. 16.

    Chen, K., Czabotar, P. E., Blewitt, M. E. & Murphy, J. M. The hinge domain of the epigenetic repressor Smchd1 adopts an unconventional homodimeric configuration. Biochem. J. 473, 733–742 (2016).

  17. 17.

    Chen, K. et al. Genome-wide binding and mechanistic analyses of Smchd1-mediated epigenetic regulation. Proc. Natl Acad. Sci. USA 112, E3535–E3544 (2015).

  18. 18.

    Brideau, N. J. et al. Independent mechanisms target SMCHD1 to trimethylated histone H3 lysine 9-modified chromatin and the inactive X chromosome. Mol. Cell. Biol. 35, 4053–4068 (2015).

  19. 19.

    Chen, K. et al. The epigenetic regulator Smchd1 contains a functional GHKL-type ATPase domain. Biochem. J. 473, 1733–1744 (2016).

  20. 20.

    Blewitt, M. E. et al. An N-ethyl-N-nitrosourea screen for genes involved in variegation in the mouse. Proc. Natl Acad. Sci. USA 102, 7629–7634 (2005).

  21. 21.

    Lemmers, R. J. L. F. et al. Digenic inheritance of an SMCHD1 mutation and an FSHD-permissive D4Z4 allele causes facioscapulohumeral muscular dystrophy type 2. Nat. Genet. 44, 1370–1374 (2012).

  22. 22.

    Gendrel, A. V. et al. Smchd1-dependent and -independent pathways determine developmental dynamics of CpG island methylation on the inactive X chromosome. Dev. Cell 23, 265–279 (2012).

  23. 23.

    Gendrel, A.-V. et al. Epigenetic functions of smchd1 repress gene clusters on the inactive X chromosome and on autosomes. Mol. Cell. Biol. 33, 3150–3165 (2013).

  24. 24.

    Leong, H. S. et al. Epigenetic regulator Smchd1 functions as a tumor suppressor. Cancer Res. 73, 1591–1599 (2013).

  25. 25.

    Mould, A. W. et al. Smchd1 regulates a subset of autosomal genes subject to monoallelic expression in addition to being critical for X inactivation. Epigenetics Chromatin 6, 19 (2013).

  26. 26.

    Nozawa, R.-S. et al. Human inactive X chromosome is compacted through a PRC2-independent SMCHD1–HBiX1 pathway. Nat. Struct. Mol. Biol. 20, 566–573 (2013).

  27. 27.

    Mason, A. G. et al. SMCHD1 regulates a limited set of gene clusters on autosomal chromosomes. Skelet. Muscle 7, 12 (2017).

  28. 28.

    Darrow, E. M. et al. Deletion of DXZ4 on the human inactive X chromosome alters higher-order genome architecture. Proc. Natl Acad. Sci. USA 113, E4504–E4512 (2016).

  29. 29.

    Deng, X. et al. Bipartite structure of the inactive mouse X chromosome. Genome Biol. 16, 152 (2015).

  30. 30.

    Giorgetti, L. et al. Structural organization of the inactive X chromosome in the mouse. Nature 535, 575–579 (2016).

  31. 31.

    Guo, Y. et al. CTCF/cohesin-mediated DNA looping is required for protocadherin α promoter choice. Proc. Natl Acad. Sci. USA 109, 21081–21086 (2012).

  32. 32.

    Brinkman, A. B. et al. Sequential ChIP-bisulfite sequencing enables direct genome-scale investigation of chromatin and DNA methylation cross-talk. Genome Res. 22, 1128–1138 (2012).

  33. 33.

    Saksouk, N. et al. Redundant mechanisms to form silent chromatin at pericentromeric regions rely on BEND3 and DNA methylation. Mol. Cell 56, 580–594 (2014).

  34. 34.

    Walter, M., Teissandier, A., Pérez-Palacios, R. & Bourc’his, D. An epigenetic switch ensures transposon repression upon dynamic loss of DNA methylation in embryonic stem cells. eLife 5, e11418 (2016).

  35. 35.

    de Greef, J. C. et al. Smchd1 haploinsufficiency exacerbates the phenotype of a transgenic FSHD1 mouse model. Hum. Mol. Genet. 27, 716–731 (2018).

  36. 36.

    Lun, A. T. L. & Smyth, G. K. diffHic: a Bioconductor package to detect differential genomic interactions in Hi-C data. BMC Bioinformatics 16, 258 (2015).

  37. 37.

    Serra, F. et al. Automatic analysis and 3D-modelling of Hi-C data using TADbit reveals structural features of the fly chromatin colors. PLoS Comput. Biol. 13, e1005665 (2017).

  38. 38.

    Lajoie, B. R., Dekker, J. & Kaplan, N. The Hitchhiker’s guide to Hi-C analysis: practical guidelines. Methods 72, 65–75 (2015).

  39. 39.

    Durand, N. C. et al. Juicer provides a one-click system for analyzing loop-resolution Hi-C experiments. Cell Syst. 3, 95–98 (2016).

  40. 40.

    Shen, Y. et al. A map of the cis-regulatory sequences in the mouse genome. Nature 488, 116–120 (2012).

  41. 41.

    Gouti, M. et al. In vitro generation of neuromesodermal progenitors reveals distinct roles for wnt signalling in the specification of spinal cord and paraxial mesoderm identity. PLoS Biol. 12, e1001937 (2014).

  42. 42.

    Gouti, M. et al. A gene regulatory network balances neural and mesoderm specification during vertebrate trunk development. Dev. Cell 41, 243–261.e7 (2017).

  43. 43.

    Lippmann, E. S. et al. Deterministic HOX patterning in human pluripotent stem cell-derived neuroectoderm. Stem Cell Rep. 4, 632–644 (2015).

  44. 44.

    Wong, S. F. et al. Independent regulation of vertebral number and vertebral identity by microRNA-196 paralogs. Proc. Natl Acad. Sci. USA 112, E4884–E4893 (2015).

  45. 45.

    Wellik, D. M. Hox genes and vertebrate axial pattern. Curr. Top. Dev. Biol. 88, 257–278 (2009).

  46. 46.

    Boulet, A. M. & Capecchi, M. R. Duplication of the Hoxd11 gene causes alterations in the axial and appendicular skeleton of the mouse. Dev. Biol. 249, 96–107 (2002).

  47. 47.

    Carapuço, M., Nóvoa, A., Bobola, N. & Mallo, M. Hox genes specify vertebral types in the presomitic mesoderm. Genes Dev. 19, 2116–2121 (2005).

  48. 48.

    Akasaka, T. et al. A role for mel-18, a Polycomb group-related vertebrate gene, during theanteroposterior specification of the axial skeleton. Development 122, 1513–1522 (1996).

  49. 49.

    Akasaka, T. et al. Mice doubly deficient for the Polycomb Group genes Mel18 and Bmi1 reveal synergy and requirement for maintenance but not initiation of Hox gene expression. Development 128, 1587–1597 (2001).

  50. 50.

    Suzuki, M. et al. Involvement of the Polycomb-group gene Ring1B in the specification of the anterior-posterior axis in mice. Development 129, 4171–4183 (2002).

  51. 51.

    van der Lugt, N. M. et al. Posterior transformation, neurological abnormalities, and severe hematopoietic defects in mice with a targeted deletion of the bmi-1 proto-oncogene. Genes Dev. 8, 757–769 (1994).

  52. 52.

    Royce-Tolland, M. E. et al. The A-repeat links ASF/SF2-dependent Xist RNA processing with random choice during X inactivation. Nat. Struct. Mol. Biol. 17, 948–954 (2010).

  53. 53.

    Trussart, M. et al. Defined chromosome structure in the genome-reduced bacterium Mycoplasma pneumoniae. Nat. Commun. 8, 14665 (2017).

  54. 54.

    Keniry, A. et al. Setdb1-mediated H3K9 methylation is enriched on the inactive X and plays a role in its epigenetic silencing. Epigenetics Chromatin 9, 16 (2016).

  55. 55.

    Naughton, C., Sproul, D., Hamilton, C. & Gilbert, N. Analysis of active and inactive X chromosome architecture reveals the independent organization of 30 nm and large-scale chromatin structures. Mol. Cell 40, 397–409 (2010).

  56. 56.

    Gordon, C. T. et al. De novo mutations in SMCHD1 cause Bosma arhinia microphthalmia syndrome and abrogate nasal development. Nat. Genet. 49, 249–255 (2017).

  57. 57.

    Shaw, N. D. et al. SMCHD1 mutations associated with a rare muscular dystrophy can also cause isolated arhinia and Bosma arhinia microphthalmia syndrome. Nat. Genet. 49, 238–248 (2017).

  58. 58.

    Jansz, N., Chen, K., Murphy, J. M. & Blewitt, M. E. The epigenetic regulator SMCHD1 in development and disease. Trends Genet. 33, 233–243 (2017).

  59. 59.

    Lemmers, R. J. L. F. et al. Facioscapulohumeral muscular dystrophy is uniquely associated with one of the two variants of the 4q subtelomere. Nat. Genet. 32, 235–236 (2002).

  60. 60.

    Gurzau, A. D. et al. FSHD2- and BAMS-associated mutations confer opposing effects on SMCHD1 function. J. Biol. Chem. 293, 9841–9853 (2018).

  61. 61.

    Vieux-Rochas, M., Fabre, P. J., Leleu, M., Duboule, D. & Noordermeer, D. Clustering of mammalian Hox genes with other H3K27me3 targets within an active nuclear domain. Proc. Natl Acad. Sci. USA 112, 4672–4677 (2015).

  62. 62.

    Gdula, M.R. et al. The non-canonical SMC protein SmcHD1 antagonises TAD formation on the inactive X chromosome. Preprint at https://www.biorxiv.org/content/early/2018/06/08/342147 (2018).

  63. 63.

    Wang, C. Y., Jégu, T., Chu, H. P., Oh, H. J. & Lee, J. T. SMCHD1 merges chromosome compartments and assists formation of super-structures on the inactive X. Cell 174, 406–421.e25 (2018).

  64. 64.

    Majewski, I. J. et al. Polycomb repressive complex 2 (PRC2) restricts hematopoietic stem cell activity. PLoS Biol. 6, e93 (2008).

  65. 65.

    Chaumeil, J., Augui, S., Chow, J. C. & Heard, E. Combined immunofluorescence, RNA fluorescent in situ hybridization, and DNA fluorescent in situ hybridization to study chromatin changes, transcriptional activity, nuclear organization, and X-chromosome inactivation. Methods Mol. Biol. 463, 297–308 (2008).

  66. 66.

    Schindelin, J. et al. Fiji: an open-source platform for biological-image analysis. Nat. Methods 9, 676–682 (2012).

  67. 67.

    Gilles, J. F., Dos Santos, M., Boudier, T., Bolte, S. & Heck, N. DiAna, an ImageJ tool for object-based 3D co-localization and distance analysis. Methods 115, 55–64 (2017).

  68. 68.

    Wingett, S. et al. HiCUP: pipeline for mapping and processing Hi-C data. F1000Res. 4, 1310 (2015).

  69. 69.

    Zhang, Y. et al. Model-based analysis of ChIP-Seq (MACS). Genome Biol. 9, R137 (2008).

  70. 70.

    Imakaev, M. et al. Iterative correction of Hi-C data reveals hallmarks of chromosome organization. Nat. Methods 9, 999–1003 (2012).

  71. 71.

    Marco-Sola, S., Sammeth, M., Guigó, R. & Ribeca, P. The GEM mapper: fast, accurate and versatile alignment by filtration. Nat. Methods 9, 1185–1188 (2012).

  72. 72.

    Le Dily, F. et al. Distinct structural transitions of chromatin topological domains correlate with coordinated hormone-induced gene regulation. Genes Dev. 28, 2151–2162 (2014).

  73. 73.

    Needleman, S. B. & Wunsch, C. D. A general method applicable to the search for similarities in the amino acid sequence of two proteins. J. Mol. Biol. 48, 443–453 (1970).

  74. 74.

    Kim, D. et al. TopHat2: accurate alignment of transcriptomes in the presence of insertions, deletions and gene fusions. Genome Biol. 14, R36 (2013).

  75. 75.

    Robinson, M. D., McCarthy, D. J. & Smyth, G. K. edgeR: a Bioconductor package for differential expression analysis of digital gene expression data. Bioinformatics 26, 139–140 (2010).

  76. 76.

    Wu, H. et al. Dnmt3a-dependent nonpromoter DNA methylation facilitates transcription of neurogenic genes. Science 329, 444–448 (2010).

  77. 77.

    Nelson, J. D., Denisenko, O. & Bomsztyk, K. Protocol for the fast chromatin immunoprecipitation (ChIP) method. Nat. Protoc. 1, 179–185 (2006).

  78. 78.

    Langmead, B. & Salzberg, S. L. Fast gapped-read alignment with Bowtie 2. Nat. Methods 9, 357–359 (2012).

  79. 79.

    Lund, E., Oldenburg, A. R. & Collas, P. Enriched domain detector: a program for detection of wide genomic enrichment domains robust against local variations. Nucleic Acids Res. 42, e92 (2014).

  80. 80.

    Buenrostro, J., Wu, B., Chang, H. & Greenleaf, W. ATAC-seq: a method for assaying chromatin accessibility genome-wide. Curr. Protoc. Mol. Biol. 109, 21.29.1–21.29.9 (2015).

  81. 81.

    Li, H. et al. The Sequence Alignment/Map format and SAMtools. Bioinformatics 25, 2078–2079 (2009).

  82. 82.

    McLean, C. Y. et al. GREAT improves functional interpretation of cis-regulatory regions. Nat. Biotechnol. 28, 495–501 (2010).

  83. 83.

    Yin, D. et al. High concordance between Illumina HiSeq2500 and NextSeq500 for reduced representation bisulfite sequencing (RRBS). Genom. Data 10, 97–100 (2016).

  84. 84.

    Krueger, F. & Andrews, S. R. Bismark: a flexible aligner and methylation caller for Bisulfite-Seq applications. Bioinformatics 27, 1571–1572 (2011).

  85. 85.

    Rigueur, D. & Lyons, K. M. Whole-mount skeletal staining. Methods Mol. Biol. 1130, 113–121 (2014).

Download references

Acknowledgements

We thank H. Coughlin, R. Allan and T. Johansson for useful discussions. This work was funded by an Australian National Health and Medical Research Council grant to M.E.B., J.M.M. and M.E.R. (GNT1098290), and fellowships to J.M.M. (GNT1105754) and M.E.R. (GNT1104924). N.J. was supported by an Australian Research Training Program Fellowship. M.E.B. was supported by a Bellberry-Viertel Senior Medical Research Fellowship. The Australian Regenerative Medicine Institute is supported by grants from the State Government of Victoria and the Australian Government. This work was made possible through Victorian State Government Operational Infrastructure Support and the Australian National Health and Medical Research Council Research Institute Infrastructure Support Scheme.

Author information

Author notes

    • Heidi Bildsoe

    Present address: Department of Molecular and Translational Science, Centre for Genetic Diseases, Hudson Institute of Medical Research, Monash University, Clayton, Victoria, Australia

    • Arne W. Mould

    Present address: Mammalian Development, Sir William Dunn School of Pathology, University of Oxford, Oxford, UK

Affiliations

  1. Walter and Eliza Hall Institute of Medical Research, Parkville, Victoria, Australia

    • Natasha Jansz
    • , Andrew Keniry
    • , Marie Trussart
    • , Tamara Beck
    • , Kelsey Breslin
    • , Megan Iminitoff
    • , Matthew E. Ritchie
    • , James M. Murphy
    •  & Marnie E. Blewitt
  2. Department of Medical Biology, University of Melbourne, Parkville, Victoria, Australia

    • Natasha Jansz
    • , Andrew Keniry
    • , Marie Trussart
    • , Matthew E. Ritchie
    • , James M. Murphy
    •  & Marnie E. Blewitt
  3. EMBL Australia, Australian Regenerative Medicine Institute, Monash University, Clayton, Victoria, Australia

    • Heidi Bildsoe
    •  & Edwina McGlinn
  4. Queensland Institute of Medical Research Berghofer, Herston, Queensland, Australia

    • Ian D. Tonks
    • , Arne W. Mould
    •  & Graham F. Kay
  5. Department of Biostatistics, Johns Hopkins University, Baltimore, MD, USA

    • Peter Hickey
  6. School of Mathematics and Statistics, University of Melbourne, Parkville, Victoria, Australia

    • Matthew E. Ritchie
  7. School of Biosciences, University of Melbourne, Parkville, Victoria, Australia

    • Marnie E. Blewitt

Authors

  1. Search for Natasha Jansz in:

  2. Search for Andrew Keniry in:

  3. Search for Marie Trussart in:

  4. Search for Heidi Bildsoe in:

  5. Search for Tamara Beck in:

  6. Search for Ian D. Tonks in:

  7. Search for Arne W. Mould in:

  8. Search for Peter Hickey in:

  9. Search for Kelsey Breslin in:

  10. Search for Megan Iminitoff in:

  11. Search for Matthew E. Ritchie in:

  12. Search for Edwina McGlinn in:

  13. Search for Graham F. Kay in:

  14. Search for James M. Murphy in:

  15. Search for Marnie E. Blewitt in:

Contributions

N.J. designed experiments, performed experiments, interpreted and analyzed data, and wrote the paper. A.K. designed experiments, performed experiments, and interpreted and analyzed data. M.T., P.H. and M.E.R. contributed to bioinformatic analyses of the data. T.B., K.B. and M.I. performed experiments. H.B. designed and performed experiments. E.M. designed experiments, performed experiments, and interpreted and analyzed data. G.F.K., I.D.T. and A.W.M. designed and performed experiments. J.M.M. designed experiments, interpreted data and edited the paper. M.E.B. designed experiments, interpreted data, and wrote and edited the paper.

Competing interests

The authors declare no competing interests.

Corresponding author

Correspondence to Marnie E. Blewitt.

Integrated supplementary information

  1. Supplementary Figure 1 In situ Hi-C-library quality-control measures.

    In situ Hi-C was performed in n=3 Smchd1fl/fl and Smchd1del/del female NSC lines, one week post Cre-mediated deletion of Smchd1. a. Tapestation trace showing read-length distribution of pooled in situ HiC libraries run on a high-sensitivity D5000 Tape. Libraries were produced in the correct size range. b. These libraries were of good quality, as determined by HiCup. Pie graph from HiCup processing report representing the relative distribution of in situ HiC reads, following optimisation of restriction digest and conditions to prevent aggregation of nuclei. Valid pairs, or chimeric HiC reads, are represented in navy blue. All other colours represent a common experimental artefact. Notably the valid reads now make up 86% of the total reads. c. Multidimensional scaling (MDS) plot showing the relationship between the 3 Smchd1fl/fl and 3 Smchd1del/del in situ Hi-C samples. Samples separate by genotype on the first MDS component: Leading logFC dim1 (X axis); Leading logFC dim1: second MDS component (Y axis). d. High correlation between biological replicates indicated by a Spearman rank correlation between bins in two matrices at increasing genomic distances. e. The log interaction count (Y axis) compared to log genomic distance (X axis) shown for the 100 kb, 500 kb and 1 Mb resolution analyses. The trend line is shown for 3 different log genomic distances. These plots show the distance-dependent decay of HiC interaction frequency.

  2. Supplementary Figure 2 In situ HiC diffHiC differential interactions and compartment analysis.

    Histograms of the differential interactions (DIs) called at 100kb (a) and 1Mb (b), plotting the distance between the two genomic anchors (Bin centre, X axis) against the frequency of interactions (Y axis). c. Summary TADbit analysis using the first eigenvector to identify compartments from a correlation matrix of normalised HiC data at 1MB resolution across chromosome 11 in Smchd1fl/fl and Smchd1del/del samples, showing no significant changes in compartment assignment (90% overlap genome-wide). The first principle component of the PCA on the Hi-C correlation matrix is shown as a green line, the density as an orange line, and the blue or red block indicates A vs B compartment d-e. Distribution of strengthened (green bars) and weakened (red bars) interactions at 1 Mb (d) and 100 kb (e) resolution. Differential interactions are spread across all chromosomes, but cluster at Smchd1 binding sites such as the X chromosome.

  3. Supplementary Figure 3 Generation of Smchd1GFP knock-in allele and its use in ChIP–seq.

    a. acGFP-loxP-neomycin resistance-loxP was knocked in immediately before the stop codon of Smchd1 in 129T2/SvEms strain mESC, such that Smchd1 would be produced as a Smchd1-GFP fusion protein. Correctly targeted mESC clones were used to generate a line of mice, and the neomycin resistance cassette removed by Cre-mediated recombination. b. The Smchd1-GFP fusion protein was detected on the inactive X chromosome, as marked by H2aK119ub. GFP and H2AK119ub immunofluorescence performed in female Smchd1GFP/GFP mouse embryonic fibroblasts. c. Optimisation of the ChIP-seq sonication conditions to release the Xi from the insoluble fraction. Sonication was performed in X129Xist∆AXCastaneus NSCs. Screenshot from genome browser showing the X chromosome. Positions of genes are shown as black boxes along the top. Tracks displayed are showing normalised counts for reads that mapped to the inactive X chromosome (castaneus) minus the reads that mapped to the active X chromosome (domesticus). Height and colour of bar represents relative coverage. Sonication time is labelled on the left. As sonication time increases, reads that map to the Xi increase. Note that even after 30 minutes sonication, coverage from the Xi is still not as great as that from the Xa (relative coverage across the X chromosome is negative). Scale bar on right is read count. Data from n=1. d. Enrichment of GFP over WCE on Chromosome 2 and Chromosome 15. Height and colour of peaks indicates strength of read counts normalised to WCE; red is most enriched, blue is most depleted. HoxD locus on Chromosome 2 and HoxC locus on Chromosome 15 are marked with an asterisk. Position of MACS2-like peaks called using Seqmonk are indicated in blue.

  4. Supplementary Figure 4 Normalized HiC interaction frequencies for interactions between the HoxB cluster and other clustered genes.

    a. Normalised Hi-C interaction frequencies at 50kb, 500 kb and 1Mb resolution displayed as a heat map and rotated 45o for Chr11:96–100Mb containing the HoxB and Keratin gene clusters. Above the horizontal is the Smchd1fl/fl heatmap, below the horizontal Smchd1del/del. The colour of the contact map, from blue to red, indicates the log2(contact frequency). Dotted white/black regions indicate the region of differential interaction, which is visible in the analyses at all 3 resolutions. b. Heatmaps displayed as in a, at 100kb resolution, for the region between the Olfactory receptor cluster genes (73 Mb) and HoxB cluster (97 Mb) on chromosome 11. Between the heatmaps, genes are shown as bars (green for Olfactory receptor genes), Smchd1 ChIP-seq peaks are shown as black bars. The white boxes bound the DIs with one anchor in the HoxB cluster. The magenta boxes are all other DIs in the region.

  5. Supplementary Figure 5 Hox genes are upregulated in Smchd1-null NMPs differentiated from mESCs.

    a. Principal component analysis (PCA) of RNAseq libraries from the differentiation of Smchd1fl/fl and Smchd1del/del mESCs to NMPs. Samples separate by timepoint on the first component (45%, x-axis) and by coverage on the second (22%, y-axis). Smchd1fl/fl samples are depicted in shades of red, and Smchd1del/del samples are depicted in shades of blue. Darker colours represent early time points, whereas lighter colours represent later time points. b. Dot plots showing expression of all genes in Smchd1fl/fl (x-axis) compared with Smchd1del/del (y-axis) in log(RPM) at D1, D5, and D8 post-induction of NMP differentiation. All genes are depicted in grey. Differentially expressed genes (FDR < 0.05) are depicted in blue. Hox genes are depicted in red. Hox genes which are differentially expressed are depicted in green. Pearson’s correlation value (R) is shown on the bottom, right of each scatter plot. c. Line graphs showing Hox gene expression throughout in vitro differentiation of male Smchd1fl/fl and Smchd1del/del mESCs into NMPs, as shown in Fig. 3a. Each graph shows expression for one paralogue group. i.e. the top left graph is showing expression of the Hox1 gene in three paralogous clusters. Time in days (x axis) is plotted against log2 transformed reads per million (RPM) (y axis). Smchd1fl/fl values are plotted in blue, and Smchd1del/del in red. Circles represent genes from the HoxA cluster, squares represent HoxB, triangles represent HoxC, and upside-down triangle for those in HoxD. Solid lines depict genes that are differentially expressed (FDR < 0.05) at D8 between Smchd1fl/fl and Smchd1del/del NMP. Dotted lines indicate genes that were not called as differentially expressed using edgeR. d. Line graphs showing the downregulation of 4/5 pluripotency genes between day 1 and day 8 of the differentiation series in Smchd1fl/fl and Smchd1del/del samples.

  6. Supplementary Figure 6 Smchd1-null embryos have altered posterior Hox-gene expression and skeletal abnormalities.

    a. Images of a representative somite-matched, sex-matched pair of E9.5 embryos, Smchd1+/+ and Smchd1MommeD1/MommeD1. b. Principal component analysis (PCA) of RNA-seq libraries from the somite-matched female paired Smchd1+/+ (n=3) and Smchd1MommeD1/MommeD1 (n=4) samples. Smchd1+/+ samples are depicted in green, and Smchd1MommeD1/MommeD1 samples are depicted in blue. c. Gene expression data from somite-matched female paired Smchd1+/+ (n=3) (y-axis) against Smchd1MommeD1/MommeD1 (n=4) samples (x-axis). Genes that are differentially expressed between Smchd1+/+ and Smchd1MommeD1/MommeD1 samples (FDR<0.05) are shown in blue. The Hox genes are shown in green, X-linked DEG in red, clustered protocadherins in orange, and Smchd1 in purple. All other genes are represented in grey. Axes are mean log transformed reads per million. d-e. Representative images from n = 10 whole mount skeletons of Smchd1+/+ (d) and Smchd1MommeD1/MommeD1 (e) E17.5 embryos, stained with Alizaren red and Alician blue. Whole body image, higher magnification view of the cervical vertebrae, and higher magnification sagittal view of embryo shown in d. Smchd1MommeD1/MommeD1 embryo shows abnormal bifurcating morphology of C2, and absence of T13 rib, indicated by asterisks. Note image for C2 is not the same Smchd1MommeD1/MommeD1 embryo as full body and sagittal view. f. Summarised data from scoring of whole mount skeletons from Smchd1+/+ (n=10) and Smchd1MommeD1/MommeD1 (n=10) male embryos. Cervical, thoracic, lumbar and sacral vertebrae are indicated, with ribs shown by horizonal blue lines, sternum shown by vertical blue lines and sacrum shown by vertical red lines. Dark blue dots indicate dramatically reduced rib process. C2bif refers to a malformed C2 vertebra, where it bifurcates at the dorsal side.

  7. Supplementary Figure 7 Smchd1 deletion does not alter the compaction accessibility of the inactive X chromosome but does result in enriched H3K27me3 on the Xi.

    a. Normalised Hi-C interaction frequencies at 100 kb resolution, generated from 200 million reads randomly subsampled from libraries at each condition, displayed as a heat map for the X chromosome in Smchd1fl/fl (a) and Smchd1del/del (b) NSCs. The colour of the contact map, from blue to red, indicates the log2(contact frequency). c. ATAC-seq was performed in Smchd1fl/fl and Smchd1del/del NSCs (n=3 female, n=2 male), one week post Cre mediated deletion of Smchd1. Position of female specific X-linked differentially accessible peaks (DAP) from the ATAC-seq analysis called using and edgeR analysis on MACS2 peaks in red. Positions of genes are indicated in black along the top. d. Sex-specific differential analysis of the MACS2 peaks identified in all samples, found 540 female-specific DAPS genome-wide. These were regions that became more accessible in the absence of Smchd1 in females, but not in males, as shown by box plots showing the coverage at 36 differential X-linked ATAC-seq peaks in log (RPM) between female Smchd1fl/fl (F) and Smchd1del/del (D) NSCs (left, n=3), and males (right, n=2). The female specific DAPs were not enriched on the X chromosome relative to autosomes, X-linked DAPs comprising 6% of the total female specific DAP (36/540). p values shown were determined by Student’s two-tailed t test. e. Confocal imaging of Smchd1fl/fl (top) and Smchd1del/del (bottom) NSC nuclei following DNA FISH with X chromosome paint, n=23 observations per genotype, n=2 cell lines. DAPI stained DNA is shown (blue) and X chromosome paint (magenta), shown with 3D rendering performed in Imaris to measure volume. f. Widefield images of H3K27me3 immunofluorescence performed in female Smchd1fl/fl (top) and Smchd1del/del (bottom) NSCs (green). g. Confocal images with Airyscan processing of H3K27me3 immunofluorescence (red) in female Smchd1+/+ (top) and Smchd1MommeD1/MommeD1 NSCs (bottom). Dot plots of volume and mean intensity of focal H3K27me3 enrichment in 3D using Imaris. Statistical comparisons were made using a two-tailed student’s t-test, line indicates mean, error bars are SEM. Measurements from 17 observations in n=2 cell lines, p <0.01. h. Genome browser tracks for the H3K27me3 ChIP-seq from ChrX:63,350,830-67,476,550 on the Castaneus-derived Xi chromosome in Smchd1del/fl and Smchd1del/del NSCs (n=2). Genes are shown in black, MACS2-called peaks in Smchd1del/fl NSCs are shown in blue, and MACS2-called peaks in Smchd1del/del NSCs are shown in red. Scale is normalised ChIP-seq read counts, normalised for read depth using the total autosomal read count.

Supplementary information

  1. Supplementary Text and Figures

    Supplementary Figures 1–7, Supplementary Note and Supplementary Table 1

  2. Reporting Summary

  3. Supplementary Dataset 1

    Uncropped Western blot images for Figure 1a

  4. Supplementary Dataset 2

    Differential interactions between Smchd1fl/fl and Smchd1del/del female NSCs, Smchd1fl/fl and Smchd1del/del male NSCs, based on diffHic analysis at 1 Mb, 500 kb and 100 kb resolutions

  5. Supplementary Dataset 3

    Strengthened and weakened TAD boundaries, TAD insulation scores, and compartment boundaries based on Tadbit analysis in Smchd1fl/fl and Smchd1del/del female NSCs

  6. Supplementary Dataset 4

    Smchd1-GFP ChIP-seq peaks

  7. Supplementary Dataset 5

    Loops defined by HiCCUPS in Smchd1fl/fl and Smchd1del/del female NSCs

  8. Supplementary Dataset 6

    Differentially expressed genes between Smchd1fl/fl and Smchd1del/del female NSCs

  9. Supplementary Dataset 7

    Differential ATAC-seq peaks between Smchd1fl/fl and Smchd1del/del NSCs

  10. Supplementary Dataset 8

    H3K27ac and H3K27me3 ChIP-seq peaks from Smchd1fl/fl and Smchd1del/del female NSCs

  11. Supplementary Dataset 9

    Differentially expressed genes between Smchd1fl/fl and Smchd1del/del male ES cells differentiating to neuromesodermal progenitors at days 1, 5 and 8 of differentiation

  12. Supplementary Dataset 10

    Differentially expressed genes between presomitic mesoderm from Smchd1+/+ and Smchd1MommeD1/MommeD1 E9.5 female embryos

  13. Supplementary Dataset 11

    Read counts for X-linked genes that mapped to the active X chromosome (genome 1) and the inactive X chromosome (genome 2) in Smchd1del/fl and Smchd1del/del female NSCs from RNAseq

  14. Supplementary Dataset 12

    H3K27me3 ChIP-seq peaks on the Xi from Smchd1del/fl and Smchd1del/del female NSCs

  15. Supplementary Dataset 13

    Read counts for RRBS data from the Xa and Xi of Smchd1del/fl and Smchd1del/del female NSCs

  16. Source Data, Figure 3f

  17. Source Data, Figure 6b, c, e, f

About this article

Publication history

Received

Accepted

Published

DOI

https://doi.org/10.1038/s41594-018-0111-z